Diseases in shrimp aquaculture have substantially reduced production and resulted in significant revenue losses (Lightner et al., “Strategies for the Control of Viral Diseases of Shrimp in the Americas,” Fish Path. 33:165–180 (1998); Dhar et al., “Isolation of Differentially Expressed Genes in White Spot Virus (WSV) Infected Shrimp (Penaeus stylirostris),” In: World Aquaculture Society, France (2000); Brock et al., “Disease Prevention and Control for Gametes and Embryos of Fish and Marine Shrimp,” Aquaculture 197:137–159 (2001)) The use of molecular biology techniques to produce pathogen-resistant strains of shrimp through genetic transformation technology is considered a highly promising strategy for control of shrimp viral disease (Mialhe et al., “Future of Biotechnology-Based Control of Disease in Marine Invertebrates,” Mol. Mar. Biol. Biotechnol. 4(4):275–83 (1995); Bachere et al., “Transgenic Crustaceans,” World Aquaculture 28(4):51–5 (1997)). In the past decade, pathogen-resistant transgenic animals and plants have been developed (Beachy, “Virus Cross-Protection in Transgenic Plants,” in D. P. S. Verma, and R. B. Goldberg, (eds.), Plant Gene Research: Temporal and Spatial Regulation of Plant Genes, New York: Springer Verlag pp. 313–327 (1998); Kim et al., “Disease Resistance in Tobacco and Tomato Plants Transformed with the Tomato Spotted Wilt Virus Nucleocapsid Gene,” Plant Dis. 78:615–21 (1993); Sin, F. Y. T., “Transgenic Fish,” Rev. Fish Biol. 7(4):417–41 (1997)), but use of such technology has only just begun for shrimp research. While methods for detecting viral disease in shrimp, including polymerase chain reaction (Dhar et al., “Detection and Quantification of Infectious Hypodermal and Hematopoietic Necrosis Virus (IHHNV) and White Spot Virus (WSV) of Shrimp by Real-Time Quantitative PCR and SYBR Chemistry,” J. Clin. Microbiol. 39:2835–2845 (2001); Tang et al., “Detection and Quantification of Infectious Hypodermal and Hematopoietic Necrosis Virus in Penaeid Shrimp by Real-Time PCR,” Dis. Aquat. Org. 44(2):79–85 (2001)), light microscopy, and transmission electron microscopy (Nunan et al., “Reverse Transcription Polymerase Chain Reaction (RT-PCR) Used for the Detection of Taura Syndrome Virus (TSV) in Experimentally Infected Shrimp,” Dis. Aquatic. Org. 34:87–91 (1998); Goarant et al., “Arbitrarily Primed PCR to Type Vibrio Spp. Pathogenic for Shrimp,” Appl. Environ. Microbiol. 65(3):1145–1151 (1999); Chen et al., “Establishment of Cell Culture Systems from Penaeid Shrimp and Their Susceptibility to White Spot Disease and Yellow Head Viruses,” Meth. in Cell Sci. 21:199–206 (1999); Toullec, “Crustacean Primary Cell Culture: a Technical Approach,” Meth. in Cell Sci. 21:193–8 (1999); Sukhumsirichart et al., “Characterization and PCR Detection of Hepatopancreatic Parvovirus (HPV) from Penaeus Monodon in Thailand,” Dis. Aquat. Org. 38:1–10 (1999), are widely used, methods for controlling viral disease in shrimp are still in development. One of the drawbacks to molecular engineering in shrimp and other crustaceans thus far has been the lack of a procedure to transform eggs or embryos with DNA that is easy, quick, highly efficient, and results in low mortality of eggs/embryos.
Three common methods of vector-expression for foreign nucleic acid delivery are electroporation (Muller et al., “Introducing Foreign Genes Into Fish Eggs With Electroporated Sperm as a Carrier,” Mol. Mar. Biol. Biotechnol. 1:276–281 (1992); Powers et al., “Electroporation: a Method for Transferring Genes Into the Gametes of Zebra Fish (Brachydanio rerio), Channel Catfish (Ictalurus punctatus), and Common Carp (Cyprimus carpio),” Mol. Mar. Biol. Biotechnol. 1:301–308 (1992); Sin et al., “Gene Transfer in Chinook Salmon by Electroporating Sperm in the Presence of PRSV-lacZ DNA,” Aquaculture 117:57–69 (1993); Powers et al., “Electroporation as an Effective Means of Introducing DNA Into Abalone (Haliotis rufescens) Embryos,” Mol. Mar. Biol. Biotechnol. 4(4):369–375 (1995); Tsai et al., “Sperm as a Carrier to Introduce an Exogenous DNA Fragment Into the Oocyte of Japanese Abalone (Haliotis divorsicolor suportexta),” Transgenic Res. 6(1):85–95 (1997); Fraga et al., “Introducing Antisense Oligonucleotides into Paramecium via Electroporation,” J. Eukaryot. Microbiol. 45(6):582–8 (1998); Preston et al., “Delivery of DNA to Early Embryos of the Kuruma Prawn, Penaeus japonicus,” Aquaculture 181:225–234 (2000)), ballistic bombardment (Zelenin et al., “The Delivery of Foreign Genes Into Fertilized Eggs Using High-Velocity Microprojectiles,” FEBS Lett. 287(1–2):118–120 (1991); Akasaka et al., “Introduction of DNA Into Sea Urchin Eggs by Particle Gun,” Mol. Mar. Biol. Biotechnol. 4(3):255–261 (1995); Gendreau et al., “Transient Expression of a Luciferase Reporter Gene After Ballistic Introduction Into Artemia Franciscana (Crustacea) Embryos,” Aquaculture 133:199–205 (1995); Baum et al., “Improved Ballistic Transient Transformation Conditions for Tomato Fruit Allow Identification of Organ-Specific Contributions of 1-Box and G-Box to the RBCS2 Promoter Activity,” Plant J. 12(2):463–9 (1997); Udvardi et al., “Uptake of Exogenous DNA Via the Skin,” J. Mol. Med. 77(10):744–50 (1999)), and microinjection (Udvardi et al., “Uptake of Exogenous DNA Via the Skin,” J. Mol. Med. 77(10):744–50 (1999); Penman et al., “Patterns of Transgene Inheritance in Rainbow Trout (Oncorhynchus Mykiss),” Mol Reprod. Dev. 30:201–206 (1991); Damen et al., “Transcriptional Regulation of Tubulin Gene Expression in Differentiating Trochoblasts During Early Development of Patella Vulgata,” Development 120:2835–2845 (1994); Gaiano et al., “Highly Efficient Germ-Line Transmission of Proviral Insertions,” Proc. Natl. Acad. Sci. USA 93:7777–7782 (1996); Cadoret et al., “Microinjection of Bivalve Eggs: Application in Genetics,” Mol. Mar. Biol. Biotechnol. 6(1):7277 (1997); Li et al., “Transfer of Foreign Gene to Giant Freshwater Prawn (Macrobrachium rosenbergii) by Spermatophore-Microinjection,” Mol. Reprod. Dev. 56(2): 149–54 (2000)). Among these three methods, microinjection is considered to be the most tedious, but most efficient, method for transferring foreign nucleic acid into marine and fresh water species. It allows precision in delivery of exogenous nucleic acid and increases the chances that a treated egg will be transformed. The introduced nucleic acid is ultimately integrated into the chromosomes of the microinjected organism. Preston et al., “Delivery of DNA to Early Embryos of the Kuruma Prawn, Penaeus japonicus,” Aquaculture 181:225–234 (2000), examined the relative efficiency of microinjection, electroporation, and particle bombardment for introducing nucleic acid into the embryos of the Kuruma prawn, Litopenaeus japonicus and found that microinjection is the most reliable technique but very time consuming. Electroporation is a desirable method for large scale gene transfer, however, host mortality tends to be high. An alternative non-surgical technique (e.g., spermatophore-microinjection), can be used as the delivery system, which provides somewhat better mortality. However, none of these methods of gene transfer is suitable to treat large numbers of fertilized shrimp eggs at one time, and most importantly, none of these methods raise the potential transformed shrimp into the mature stage.
Transgenic techniques provide a potential tool in producing shrimp capable of combating diseases and, subsequently, improving aquaculture production. However, at present, transgenic shrimp studies suffer from the lack of availability of suitably efficient methods for the introduction of foreign DNA into the very fragile shrimp zygotes. What is needed is a method of in vivo DNA delivery into the eggs of shrimp and other that provides improved ease of use, improved efficiency of transformation, and improved mortality rates over the existing methods.
The present invention is directed to overcoming these and other deficiencies in the art.